where r = RU / RUmax,K1 and K2 are the variableparameters to fit, RU is the response at the steady-state level, RUmax is the maximum response for binding one molecule per binding site, Cfree is the concentration of the compound in solution and K1 and K2 are the macroscopic binding constants. K2 is zero for a single-site binding model. The RUmax was determined as previously described (40) from the DNA molecular weight, amount of DNA on the flow cell, the compound molecular weight and the refractiveindexgradientratio of the compound and DNA. The Kvalues are determined for each set of sensorgrams by nonlinear least square fitting of rversusCfreeplots for compound bound to each DNA.

The structure of a complex between compound 3 and the dodecamer duplex DNA sequence d(CGCGAATTCGCG)2 was constructed, starting from the known structure of the dodecanucleotide complex with compound 1 (seeabove). The geometry and orientation of the bis-benzimidazole group were leftexactly as they were in the X-ray crystal structure. Instead the terminal dimethylaminopropyloxy side chains of 1 were mutated into the two piperidinopropyl groups of 3. The solvated complex was then subjected to 3 ps molecular dynamics at 300 K followed by 200 steps of conjugate gradient minimization with a 50 kcal/mol-residue harmonic restraint on the DNA and on the bis-benzimidazole centralmoiety. The complex was then neutralized, solvated and equilibrated using the same protocols as described above for 1. Finally, a dynamics production run of 1000 ps (1 fs step) was carried out at 300 K. Relative binding energies were calculated (42) from the structures averaged over the production runs, which were then subjected to molecular mechanicsminimizations to convergence.

Sequence selective binding. The gels show DNase I footprinting with (A) 117mer and (B) 178mer PvuII–EcoRI restriction fragments cut from the plasmids pBS and pKS, respectively. In both cases, the DNA was labeled at the EcoRI site with [α-32P]dATP in the presence of AMV reverse transcriptase. The products of nuclease digestion were resolved on an 8% polyacrylamide gel containing 7 M urea. Control tracks (Cont) contained no drug. The concentration (µM) of the drug is shown at the top of the appropriate gel lanes. Tracks labeled ‘G’ represent dimethylsulfate-piperidine markers specific for guanines. Numbers on the side of the gels refer to the standard numbering scheme for the nucleotide sequence of the DNA fragment.

Figure 3

Differential cleavage plots comparing the susceptibility of (A) the 117mer and (B) the 265mer pBS restriction fragments to DNase I cutting in the presence of the bis-benzimidazole compounds (2 µM each). Negative values correspond to a ligand-protected site and positive values represent enhanced cleavage. Vertical scales are in units of ln(fa) – ln(fc), where fa is the fractional cleavage at any bond in the presence of the drug and fc is the fractional cleavage of the same bond in the control, given closely similar extents of overall digestion. Each line drawn represents a 3-bond running average of individual data points, calculated by averaging the value of ln(fa) – ln(fc) at any bond with those of its two nearest neighbors. Only the region of the restriction fragment analyzed by densitometry is shown. Black boxes indicate the positions of inhibition of DNase I cutting in the presence of the drugs.

Similarconclusions can be made from the experiments performed with the 178 bp fragment. As shown in the gel in Figure 2B, compound 1 binds very strongly to the AATT site around position 53, whereas this site is filledmuchmoreweakly by compounds 2 and 3. A detailed comparison of the binding to DNA of compound 1 and Hoechst 33258 is presented in the gel shown in Figure 4, and the corresponding cleavage plots in Figure 5. Although from the gel the footprinting patternsappearvisually very similar for the two drugs, the densitometric analysis reveals interestinglocaldifferences. At 1 µM, compound 1 and Hoechst 33258 bind equallywell to the 5′-AATT, TAATA and TAAAA sites at positions 53, 75 and 100, respectively. The fourth binding site identified around position 120 corresponds to the sequence 5′-TTTT to which compound 1 appears to bind more strongly than the Hoechst dye (Fig. 5A). This is consistent with what was observed with the other DNA fragments. Indeed, the same sequence occurred in the 117 bp fragment around position 64 and at that site we also detected a higher site occupancy for compound 1 versus Hoechst 33258 (Fig. 3A). At a slightly higher concentration, 3 µM, the magnitude of the footprint at the 5′-TTTT site is identical for the two drugs, as is the case for the 5′-AATT and 5′-TAAAA sites (Fig. 5B). But at this concentration a new footprint was detected at the sequence TATA around position 63 (openrectangle in Fig. 5B) for compound 2 but not for Hoechst 33258. Similarly, a weak footprint appearedspecifically with the symmetric head-to-head bis-benzimidazole derivatives around position 86 at a mixed TGAG sequence flanked by GC tracts, whereas no effect was seen with the head-to-tail compound Hoechst 33258 (open rectangle in Fig. 5B). The two sequences (purely GC) flanking this new binding site become more susceptible to attack by DNase I than in the control in the presence of compound 1. The cleavage enhancement at these GC sites may be attributable to drug-induced perturbations of the double helical structure of DNA. Compound 1 strongly discriminates between runs of guanines and/or cytosines. From the footprinting experiments, we concludedunambiguously that the binding of the bis-benzimidazole derivative 1 to AT sequences was much favored over binding to GC or mixed sequences.

Figure 4

DNase I footprinting of compound 1 and Hoechst 33258 on the 174 bp PvuII–EcoRI DNA fragment from plasmid pKS. Numbers on the side of the gel refer to the standard numbering scheme for the nucleotide sequence of the DNA fragment, as indicated in Figure 5. Other details as for Figure 2.

Figure 5

Differential cleavage plots comparing the susceptibility of the 174mer pKS DNA fragment to DNase I cutting in the presence of compound 1 or Hoechst 33258 at (A) 1 or (B) 3 µM. Filled boxes indicate the positions of binding sites common to the two compounds. Open boxes refer to the position of binding sites specific to compound 1.

BIAcore surface plasmon resonance experiments

Binding studies with DNA hairpin–duplex oligomers were conducted with a BIAcore 3000 SPR instrument. The neutral compound 2 did not have sufficient solubility in buffer for BIAcore experiments. To compare the complexes of the compounds with A·T and G·C base pair sequences, the association of the compounds with two DNA oligomer hairpin–duplexes with alternating A·T or G·C base pairs was determined as described in the Materials and Methods. Sensorgrams at increasing compound concentrations for interaction with the two DNAs revealsignificant differences between the DNA complexes (Fig. 6). Both compounds bind strongly to the AT DNA duplex and reachsaturation in the concentration range used in these experiments (0–500 nM) as can be seen from the sensorgrams. With the GC sequence, however, no binding of compound 1 can be detected over the same concentration range, while quite significant binding of compound 3 is observed. The binding of compound 3 to the GC sequence is, however, weaker than its interaction with the AT DNA. The sensorgram results were fit in the steady-state region as described in the Materials and Methods, and binding constants are collected in Table1. As expected from observation of the sensorgrams, the binding constants for binding to the AT DNA are large (4–7 × 107) and similar for the two compounds. The binding constant for compound 1 with the GC DNA must be <106since no binding is detected, while the binding of compound 3 to the GC DNA has a binding constant of 1.9 × 106. The GC binding constant for compound 3 is ∼10 times lower than the AT binding constant for the same compound (Table 1). The AATT sequence also has a strong footprint with compound 1. BIAcore results for compounds 1 and 3 binding to an AATT DNA sequence are shown in Figure 7. Again, the binding is strong and the magnitude of the binding constants is similar for the two compounds. As with the alternating sequence AT DNA, compound 1 binds reproducibly more strongly to the AATT DNA than compound 3 (9.4 versus 2.1 × 107 by steady-state analysis; Table 1).

Figure 6

BIAcore SPR sensorgrams for the interaction of compounds 1 and 3 with the alternating AT and GC sequence DNA hairpins in HBS buffer at 25°C. Both compounds bind strongly to the AT sequence and reach saturation of the DNA in the concentration range of this experiment, 1–500 nM compound. In the same concentration range, no binding of compound 1 to the GC sequence is observed while significant binding of compound 3 can be detected. Fitting of results from these and additional experiments in the steady-state region provided data for determination of compound binding constants, as described in the Materials and Methods, and these are collected in Table 1.

Figure 7

BIAcore SPR sensorgrams for the complexes of compounds 1 and 3 with the AATT DNA minor groove in HBS buffer at 25°C. Both compounds bind strongly to the AATT sequence and reach saturation at concentrations below 500 nM. Global fitting of the curves to obtain association and dissociation kinetics constants was done with BIA Evaluation software and a single site interaction model (Materials and Methods). The best-fit lines through each experimental plot are also shown in the Figure and as can be seen, the global, single-site model provides excellent fits to all of the experimental curves. Similar fits with the other DNA samples provided the kinetics constants in Table 1.

Table 1.

Keqb (×10–7)

kac (×10–5)

kdc (×103)

ka / kd (×10–7)

1-AATT

9.4

0.61

1.7

3.6

1-AT

6.3

1.1

1.3

8.1

1-GC

<1

3-AATT

2.1

3.0

12

2.6

3-AT

2.0

3.4

5.1

6.6

3-GC

0.18

0.47

25

0.20

Equilibrium and kinetic constants for the interaction of compounds 1 and 3 with different sequence DNAs by SPR analysisa

Visual observation of the binding sensorgrams for the two compounds interacting with the AT DNA sequence indicated that eventhough the binding affinities were similar for the compounds at the AT sites, the rates of binding were different. Compound 1 has both slower association and slower dissociation kinetics than compound 3. Global kinetics fits to the sensorgrams for compounds 1 and 3 with the AATT sequence, where there is one specific binding site, are shown in Figure 7 and the kinetics results are also collected in Table 1. The sensorgrams for compound 1 decrease slightly with time at the highest concentrations used in the experiments, but deletion of these curves with subsequent fitting suggests that the decrease does not cause a large error in fitting in this case. With the AATT sequence the curves are fit quite well with a model having one binding site per DNA hairpin and the residuals are generally small. With the alternating AT and GC DNA samples, the kinetics results suggestsome weaker secondary sites of compound binding are present. The results in Table 1 are for binding to the strong primary binding site. With the AATT DNA sequence where the most accurate fitting could be done, both the on and off rates for compound 1 are five to seven times lower than for compound 3. Binding constants determined by the ratio of ka/kd are not as accurate as with the steady-state fitting method but are in the same range as obtained with the steady-state fits (Table 1). With the GC sequence, kinetics constants could only be determined with compound 3 due to the very weak binding of compound 1 to the GC DNA. As can be seen from the results in Table 1, the lower binding affinity of compound 3 for the GC relative to the AT sequence DNA is due to a combination of lower association and larger dissociation rate constants. The equilibrium constants calculated by steady-state and kinetics methods are quite close for the GC DNA sequence.

Molecular modeling

The weak footprints obtained with the neutral compound 2 could be predicted, but similar results were expected both in terms of footprints and binding affinity for AATT sequences for the two doubly charged ligands 1 and 3. Compound 3 hadoriginally been designed to increase the binding affinity of compound 1 since it has the two terminal methyl groups replaced with a cyclic group having a larger vanderWaals surface. In order to rationalize the experimental data obtained from DNase I footprinting and BIAcore SPR experiments, molecular mechanics and dynamics simulations have been performed for complexes 1 and 3. Both complexes have been studied in an explicit water solvated environment using standard AMBER protocols for dynamic simulations. The structures of both compound 1 and 3 stayedstablethroughout all the 1000 ps of dynamic simulation and theirshapesadapted very well to the minor groove curvature. The hydrogen bond donors and the aromaticsystems of the two benzimidazole subunitsformed stable non-covalent hydrogen bond interactions with the four AT base pairs in the center of the sequence, whereas differences were observed in the way that the side chains of the two drugs interacted with the GC base pairs adjacent to the AATT central core. A well localized and relatively immobile water molecule mediated the non-covalent interaction between each side chain of compound 1 and the DNA, through the formation of a triad of hydrogen bondsinvolving the phenoxy oxygen atom in the ligand molecule and the O2 and N3 hydrogen bond acceptoratoms in the cytosine and adenine bases (Table 2 and Fig. 8). There are no contacts <3.6 Å between the water molecules and ligand side chain carbon atoms. The hydrogen bond interactions would be expected to significantly enhance the binding affinity between compound 1 and the DNA. In compound 3, on the other hand, the replacement of the phenoxy oxygen in each side chain by a carbon atom confersgreaterhydrophobiccharacter to the side chains and removes the possibility of water-mediated hydrogen bonding to bases. During the dynamics simulation only one water molecule was observed in the region between the drug and the DNA (Fig. 9). This single water molecule created stable hydrogen bonds with O2 of a cytosine and N3 of a guanine, but no specific interactions with the ligand were observed.

Figure 8

Plots of the averaged MD structure of the complex of compound 1 with d(CGCGAATTCGCG)2, showing the water molecule bridging between the ligand and base edges. Hydrogen bonds are shown as dashed lines.

Figure 9

Plots of the averaged MD structure of the complex of compound 3 with d(CGCGAATTCGCG)2, showing the sole water molecule in the vicinity of the ligand–DNA interface.

Table 2.

Distances (Å)

Angles (°)

O1drug

O2drug

O2

N3

Od–Ow–O2

Od–Ow–N3

O1w

3.19

–

3.36

2.93

84

132

O2w

–

3.16

3.31

2.95

84

138

Geometric features of the hydrogen bonding arrangement around the water molecule in the complex with ligand 1, taken from the averaged structure, as shown in Figure 8

The relative binding energy of compound 1 to the d(CGCGAATTCGCG)2 structure was calculated to be –112 kcal/mol, and that of compound 3 to be –92 kcal/mol. We suggest that the tight hydrogen bond network observed in the complex with compound 1, compared with its absence in the complex with compound 3, is a majorfactor in the lower binding affinity of compound 3 and its weak DNA footprint. Although such calculations are necessarilyapproximate, it is reassuring that the difference in relative binding energies, of 20 kcal/mol, is what would be expected for the difference of six hydrogen bonds between the complexes. The terminal piperidine groups in compound 3, even though they have a larger van der Waals interaction surface than the methyl groups in compound 1, are not fullyembedded in the groove, and thus do not offset the hydrogen bonding contributions made by the latter. Figure 10 shows that the absence of the bridging water molecule in the complex with compound 3 results in the appearance of a small void between the ligand and minor-groove surface.

Figure 10

Plots of the solvent-accessible surface of the d(CGCGAATTCGCG)2 duplex together with the van der Waals surface of each ligand, viewed down one end of the minor groove for each averaged structure, (A) showing the complex with ligand 1, with the bound water molecule shown in green, and (B) showing the complex with ligand 3. A small void in the groove is apparent between this ligand and the DNA.

14.CarrondoM.A.A. de C.T., Coll,M., Aymami,J., Wang,A.H.-J., Van der Marel,G.A., Van Boom,J.H. and Rich,A. (1989) Binding of a Hoechst dye to d(CGCGATATCGCG) and its influence on the conformation of the DNA fragment. Biochemistry, 28, 7849–7859.2482071

41.RyckaertJ.P., Ciccotti,G. and Berendsen,H.J.C. (1977) Numerical integration of the cartesian equations of motion of a system with constraints: Molecular dynamics of n-alkanes. J. Comput. Phys., 23, 327–341.

46.NgyenB., Lee,M.P.H., Hamelberg,D., Joubert,A., Bailly,C., Brun,R., Neidle,S. and Wilson,W.D. (2002) Strong binding in the DNA minor groove by an aromatic diamidine with a shape that does not match the curvature of the groove. J. Am. Chem. Soc., 124, 13680–13681.12431090

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